Enzymes are the workhorses of a cell that affect virtually every biological process that characterizes a living organism. They catalyze chemical reactions with remarkable specificity and rate enhancements. The awesome catalytic power and versatility of enzymes has long been recognized and enzymes have proved to be very useful outside the living system as well. Enzymes today have widespread application in industry and are seen as environment friendly alternatives to chemical reagents because enzymatic reactions require milder conditions and tend to be cleaner with lesser byproduct and waste generation. Enzymes are being used in numerous new applications in the food, feed, agriculture, paper, leather, and textiles industries, resulting in significant cost reductions and environment-friendly operations.
Enzymes have evolved to function best under the physiological conditions of the parent organism. In vitro applications often call upon enzymes to work under non-physiological conditions or to perform functions they have not evolved for. For example, enzymes may have to catalyze reactions involving novel substrates; they may have to work under extreme conditions of salt, temperature, pH etc., or in the presence of potentially inhibiting or denaturing chemicals. Such applications have brought to light the severe disabilities of enzymes to function as industrial catalysts. In order to extract optimum performance from enzymes in the test-tube and in industrial reactors, these biocatalysts need to be tailored to suit specific applications.
The commercial success of these enzymes can be attributed to their ease of use. In addition to functioning at high temperatures, thermostable enzymes generally posses an increased shelf life which markedly improves handling conditions. If enzymes are to play a significant role in large scale processing of chemicals, they must be able to endure the harsh conditions associated with these processes. Thermostable enzymes are easier to handle, last longer, and given the proper immobilization support should be reusable for multiple applications.
In obtaining thermostable enzymes the conventional approach is to screen the microbial collections collected from extremephillic environments (Karsheroff and Ladenstein, 2001). The promising candidate enzymes are further investigated for suitability for a specific process. For example, for applications requiring thermostable or salt stable enzymes, enzymes from thermophilic or halophilic organisms were used, respectively.
However, such an approach severely restricted the use of enzymes because enzymes for all applications cannot be found in nature. There may not be a natural enzyme for many kinds of transformations. Moreover, enzyme usage is often restricted by undesirable properties of enzymes like product inhibition, low stability etc. Very often an enzyme is required to have a combination of several properties that may be impossible to find in a natural enzyme. Another approach to obtain thermostable enzymes is based on the current knowledge on the protein structures (crystal structures) of homologous enzymes from mesophiles and thermophiles (Kumar et al, 2000; Lehmann et al, 1998). Such comparisons yielded information on the probable interactions that enhance thermostability. Using such information efforts were made to incorporate these changes in mesophillic enzymes to improve their thermostability. Such approaches have not been very successful since interactions that improve thermostability in a protein are many and each protein acquires, over evolutionary times, those interactions that are best suited for its sequence and the mileu in which it functions. Though structural determinants of protein stability have been objects of numerous studies on model proteins, no universal stabilization mechanism has yet been found (Jaenicke and Bohm, 1998). The most obvious conclusion that can be drawn from the literature is that different proteins have adapted to different thermal environments by a variety of evolutionary devices. The lack of understanding of the structural features leading to protein thermostability has been partly due to a scarcity of data because experimental studies comparing homologous proteins from psychrophilic, mesophilic and thermophilic organisms have been limited to only a few proteins. Moreover, inability to form definite rules for improving protein thermostability is due to the large number and complexity of possible contributing factors (Jaenicke and Bohm, 1998; Vogt et al. 1997a; Vogt et al., 1997b; Ladenstein and Antranikain, 1998). Based on comparisons between mesophiles and thermophiles, the main mechanisms responsible for increased thermostability have been identified as increase in the number of hydrogen bonds and salt bridges, increased core hydrophobicity, better packing efficiency, α-helix and loop stabilization and resistance to covalent destruction. Often it becomes difficult to delineate protein interactions that contribute to thermostability from other selection pressures such as salt, pH etc.
Other strategies adapted to increase the thermostability was based on the observation that immobilized enzymes acquire thermostability to some extent (Reetz et al., 1995). Hence, several solid supports were tried to immobilize proteins. And also recent observations made with enzymes in organic solvents indicated that in organic solvents enzymes acquire thermostability (Plou and Ballesteros, 1999). The advent of recombinant DNA techniques has greatly facilitated protein engineering by allowing facile mutagenesis and production of proteins.
The term protein thermostability refers to the preservation of the unique chemical and spatial structure of a polypeptide chain under extremes of temperature conditions (Jaenicke and Bohm, 1998). In general, the higher the temperature to which the enzyme is exposed, the shorter the half-life of the enzyme (i.e., the shorter the enzyme retains its activity). Similarly, the greater levels of organic solvent to which said enzymes are exposed, the shorter the half-life of the enzyme. The phrase “catalytic activity” or simply “activity,” means an increase in the k.sub.cat or a decrease in the K.sub.M for a given substrate, reflected in an increase in the k.sub.catt/K.sub.M ratio. The structural basis of protein thermostability has been an actively pursued area of research for at least two decades (Argos et al., 1979). However, enzymes lifted out of the context of living organisms do not always function as well as they do in their natural milieu. For example, they have optimum activity at the physiological temperature of the organism and tend to denature at higher temperatures leading to drop in activity. Thermostable enzymes are important as they can be used at high temperatures and harsher conditions required in industrial contexts. Also they generally have higher storage stabilities and bring down costs by obliviating the need for low temperature storage and decreasing the loss due to denaturation on storage and handling. Moreover reactions carried out at higher temperatures generally proceed at higher rates further bringing down operation times.
In view of the environmental safety reasons, there is a constant pressure to reduce the use of environmentally polluting processes in industry. Enzymes are increasingly used to replace chemical processes in leather, food, and pharmaceutical industries. Comparison of protein structures from extremeophiles demonstrated that protein structural plasticity is enormous and is resident in the primary structure. This lent considerable support to strategies that alter the primary structure of the proteins at the genetic level and screen for the variants with special properties such as thermostability. The tremendous success in handling the genes and developing protocols to alter it at will, has allowed to evolve proteins with special functions. The strategy relies in generating variation in gene sequences by molecular biology methods and screening the variants by expressing them and screening the mutant population (Arnold, 1999; Stemmer, 1994; Ostermeier et al., 1999). The screening protocols are based on the property of interest, e.g., activity at high temperature or activity in the presence of organic solvents. The present invention encompasses methods for generating variation in gene sequences, protocols for screening the enzymes with higher thermostability and also protocols for sequencing and expression.
Lipases (triacylglycerol acylhydrolases, E.C. 3.1.1.3) are water-soluble enzymes that catalyze the hydrolysis of ester bonds in triacylglycerols and often also exhibit phospholipase, cutinase and amidase activities (Woolley and Petersen, 1994). They are used for the production of detergents, pharmaceuticals, perfumes, flavour enhancers and texturising agents in consmetic products. Lipases are crucial for the production of a wide variety of foods, especially for products from milk, fat and oil. Lipases are ubiquitous enzymes of considerable physiological significance and industrial potential. Lipases catalyze the hydrolysis of triacylglycerols to glycerol and free fatty acids. In contrast to esterases, lipases are activated only when adsorbed to an oil-water interface and do not hydrolyze dissolved substrates in the bulk fluid. A true lipase will split emulsified esters of glycerine and long-chain fatty acids such as triolein and tripalmitin. Lipases are serine hydrolases. Commercially useful lipases are usually obtained from microorganisms that produce a wide variety of extracellular lipases (Jaeger et al., 1999). Many lipases are active in organic solvents where they catalyze a number of useful reactions including esterification transesterification, regioselective acylation of glycols and menthols, and synthesis of peptides and other chemicals. An increasing number of lipases with suitable properties are becoming available and efforts are underway to commercialize biotransformation and syntheses based on lipases (Schmid and Verger, 1998). Enzyme sales for use in washing powders still remain the single biggest market for industrial enzymes. The major commercial application for hydrolytic lipases is their use in laundry detergents. Detergent enzymes make up nearly 32% of the total lipase sales. Lipase for use in detergents needs to be thermostable and remain active in the alkaline environment of a typical machine wash. An estimated 1000 tons of lipases are added to approximately 13 billion tons of detergents produced each year Because of their ability to hydrolyzes fats, lipases find a major use as additives in industrial laundry and household detergents. Detergent lipases are especially selected to meet the following requirements: (1) a low substrate specificity, i.e., an ability to hydrolyze fats of various compositions; (2) ability to withstand relatively harsh washing conditions (pH 10-11, −60° C.); (3) ability to withstand damaging surfactants and enzymes [e.g., linear alkyl benzene sulfonates (LAS) and proteases], which are important ingredients of many detergent formulations. Lipases with the desired properties are obtained through a combination of continuous screening (Jaeger and Reez, 1998; Wang et al.; Rubin and Dennis, 1997) and protein engineering (Kazlauskas and Bornscheuer, 1998). In 1994, Novo Nordisk introduced the first commercial recombinant lipase ‘Lipolase,’ which originated from the fungus Thermomyces lanuginosus and was expressed in Aspergillus oryzae. In 1995, two bacterial lipases were introduced—‘Lumafast’ from Pseudomonas mendocina and ‘Lipomax’ from P. alcaligenes—by Genencor International (Jaeger et al., 1999al). According to a report an alkaline lipase, produced by P. alcaligenes M-1, which was well suited to removing fatty stains under conditions of a modern machine wash. The patent literature contains examples of many microbial lipases that are said to be suitable for use in detergents (Gerritse et al., 1998).
Lipase-producing microorganisms include bacteria, fungi, yeasts, and actinomyces. Bacillus subtilis is a Gram-positive, aerobic, spore-forming bacterium that has generated substantial commercial interest because of its highly efficient protein secretion system Though extracellular lipolytic activity of B. subtlis was observed as early as in 1979 (Bycroft and Byng, 1992), molecular research started in 1992 when a lipase gene, lipA, was cloned and sequenced (Dartois et al., 1992). Subsequently the lipase was overexpressed, purified and characterized (Leuisse et al., 1993). Later, a second gene, lipB, was found that is 68% identical with lipA at the nucleic acid level (Eggert et al., 2000). This gene has been cloned and the protein overexpressed, purified and characterized.
The Bacillus subtilis lipase with a molecular weight of 19,348 Da is one of the smallest lipases known. It is one of the few lipases that do not show the interfacial activation in the presence of oil-water interfaces. LipA is very tolerant to basic pH and has its optimum activity at pH10. It hydrolyses the sn-1 and sn-3 glycerol esters with both short and long chain fatty acids, showing optimum activity with C8 fatty acid chains.
Bacterial lipases are classified into eight families according to their sequence similarities, conserved sequence motifs and biological properties (Arpigny and Jaeger, 1999). The true lipases are classified in family I which contains six subfamilies. Bacillus lipases have been placed in subfamilies 4 and 5. In these two subfamilies alanine replaces the first glycine residue in the conserved G-X-S-X-G pentapeptide around the active site serine residue. Subfamily 4 consists of only three members, LipA and LipB from B. subtilis and a lipase from Bacillus pumilis, which share 74-77% sequence identity. These are the smallest lipases known and show very little sequence similarity (˜15%) with the other, much larger, Bacillus lipases that constitute subfamily 5.
The crystal structure of the B. subtilis lipase LipA reveals a globular protein with dimensions of 35×36×42 (Pouderoyen et al., 2001). The structure shows a compact domain that consists of six β-strands in a parallel β-sheet, surrounded by α-helices. There are two α-helices on one side of the α/β sheet and three on the other side. The fold of the B. subtilis lipase resembles that of the core of the α/β hydrolase fold enzymes. The B. subtilis lipase lacks the first two strands of the canonical α/β hydrolase fold and the helix αD is replaced by a small 310 helix. The helix αE is exceptionally small, with only one helical turn, and several α-helices start or terminate with 310 helical turns. Due to these structural features, its small size and absence of a lid domain, the B. subtilis lipase is considered a minimal α/β hydrolase fold enzyme.